This work was supported by National Eye Institute grants EY02665 and EY031706 and International Retinal Research Foundation to Dr. Vinberg, National Institutes of Health Core Grant (EY014800), and an Unrestricted Grant from Research to Prevent Blindness, New York, NY, to the Department of Ophthalmology &#38; Visual Sciences, University of Utah. Dr. Frans Vinberg is also a recipient of a Research to Prevent Blindness/Dr. H. James and Carole Free Career Development Award, and Dr. Silke Becker of an ARVO EyeFind grant. We thank Dr. Anne Hanneken from The Scripps Research Institute for providing the donor eye used for recordings shown in Figure 2E.

All experiments using mice were conducted in accordance with the NIH Guide for the Care and Use of Laboratory Animals and were approved by the Institutional Animal Studies Committee at the University of Utah. Pig eyes for demonstration of this video were obtained postmortem from the slaughterhouse (Sustainable Swine Resources, Johnsonville). Eyes were obtained from human donors after brain or cardiac death with consent for research use through the Utah Lions Eye Bank, the San Diego Eye Bank or Lifesharing, which are fully accredited by the FDA, the Association of Organ Procurement Organizations (AOPO) and the Eye Banks of America Association. The use of human donor eyes had exempt status at the University of Utah (IRB no. 00106658) and ScrippsHealth IRB (IRB no. 16-6781).
1. Setting up the ex vivo ERG
<ol>
	<li>To convert a multielectrode array setup, connect the ex vivo ERG specimen holder via a head stage to a differential amplifier, which is plugged into the analog input of the interface board of the multielectrode array system. Use the recording software for the multielectrode array to record and store the input data from the ex vivo ERG. Set the gain of the differential amplifier to 100 and add an additional 10x voltage amplification depending on the digitizer specifications. Set the low pass filter to 100 Hz.</li>
	<li>To convert a patch clamp setup, connect the ex vivo ERG specimen holder via a head stage to a differential amplifier, which is connected to the head stage of the patch clamp amplifier. Use the patch clamp system software and digitizer to record and store the input data from the ex vivo ERG. Set the gain of the differential amplifier to 100 and apply an additional 10x voltage amplification via the patch clamp head stage. Set the low pass filter to 100 Hz.</li>
	<li>Connect LEDs with the appropriate wavelengths (e.g., approximately 530 nm to elicit rod photoreceptor and ON-bipolar cell responses) to the microscope. Control the LEDs with recording software that enables the triggering of light stimuli. To control light stimuli, use an LED driver controlled by analog outputs from a digitizer.</li>
	<li>Calibrate the light output of the LEDs at the position of the retina in the specimen holder using a photodiode. If necessary, insert neutral density filters&#160;into the light path to dim the light intensity.</li>
	<li>Use a commercially available or custom built ex vivo ERG specimen holder. 
	NOTE: Drawings for CNC machining from polycarbonate can be obtained from the authors upon request.</li>
	<li>To prepare the electrode, insert an Ag/AgCl pellet electrode into a threaded luer connector. Fill the inside of the luer connector with hot glue and insert a 2 mm socket into the hot glue from the non-threaded side. Solder a silver wire of the EP1 electrode to the 2 mm socket. Screw the finished electrode, with an o-ring on the thread, into the electrode ports of the ex vivo ERG specimen holder. 
	&#8203;NOTE: Electrodes can be used for a long time. If the pellet surface accumulates dirt and/or gets dark (this can cause high offset voltage and/or electrical drift), it can be &#34;polished&#34; using fine sandpaper.</li>
	<li>At least 1 day prior to experimentation, glue filter papers onto the domes of the ex vivo ERG specimen holder using epoxy glue, ensuring that the glue does not obstruct the electrode channel (see 3 for video).</li>
</ol>
2.&#160; Animal preparation
<ol>
	<li>Dark adapt animals for at least 6 h or overnight.</li>
</ol>
3. Equipment preparation
<ol>
	<li>Fill the perfusion line of the&#160;ex vivo&#160;ERG with Ames&#39; medium bubbled with 5% carbon dioxide and 95% oxygen. Connect the perfusion line to a heating element with a DC current source or heat controller near the specimen holder to warm the Ames&#39; medium, so that the retina is kept at approximately 35-38 &#176;C.</li>
	<li>Fill both halves of the&#160;ex vivo&#160;ERG specimen holder with electrode solution, seal the perfusion line with luer stoppers, connect the electrodes, and assemble the specimen holder with four screws (see&#160;3&#160;for video).</li>
	<li>Check the resistance and offset voltage between the electrodes in the assembled specimen holder by inserting the probes of the multimeter into the electrodes. 
	NOTE: If there are no blockages, and electrodes are in good condition, the resistance should be below 100 k&#937; and offset voltage below 5 mV.</li>
	<li>Connect the assembled specimen holder to the perfusion line and place onto the stage of a microscope set up to deliver light flashes. Ensure that the specimen holder and perfusion line do not contain any air bubbles.</li>
</ol>
4. Tissue preparation
<ol>
	<li>Sacrifice the animal with CO2 and immediately enucleate the eyes, or obtain large animal or human donor eyes.</li>
	<li>Clean the globe of the remaining connective tissue and extraocular muscles. Trim off the optic nerve.</li>
	<li>On filter paper, carefully place a small incision with a razor blade along the ora serrata, approximately 1 mm from the limbus in the mouse eye. Insert fine vannas scissors into the incision and cut along the limbus to remove the anterior portion of the eye with the lens.</li>
	<li>Move the eye cup into a dish containing Ames&#39; medium. Grasp the sclera with fine forceps, taking care to not touch the retina. Insert vannas scissors between the retina and the sclera and cut the sclera from the peripheral toward the central part. Take care not to touch or damage the retina.</li>
	<li>Immobilize the sclera by holding one side of the incision with vannas scissors against the bottom of the dissection dish. Grasp the sclera with forceps on the other side of the incision. Remove the sclera without touching or damaging the retina by pulling apart the sclera on either side of the incision to allow isolation of the retina with minimal damage to the tissue.</li>
	<li>Remove the anterior segment with the lens from the fellow eye and store the eye cup at room temperature in Ames&#39; solution bubbled with 5% carbon dioxide and 95% oxygen. 
	NOTE: Functional light responses from eyes stored in this way can be obtained for several hours.</li>
	<li>In large eyes, including human donor eyes, clean the globe of the remaining connective tissue and remove the anterior segment and lens, similarly to the procedure described for the mouse eye. Use a scalpel to make a cut approximately 3 mm from the limbus. Insert curved dissection scissors into the incision and cut along the limbus to remove the anterior portion of the eye with the lens. Obtain retinal specimens for ex vivo electroretinography with a 3-6 mm disposable biopsy punch.</li>
</ol>
5. Mounting the tissue on the specimen holder
<ol>
	<li>Place the lower half of the specimen holder into a large Petri dish and fill with oxygenated Ames&#39; medium so the dome of the specimen holder is just submerged.</li>
	<li>Carefully grasp the edge of the retina with fine forceps and transfer the retina onto the dome of the ex vivo specimen holder, photoreceptor side facing up.</li>
	<li>Lift the specimen holder from the Ames&#39; solution, taking care that the retina stays in place.</li>
	<li>Completely dry the plate of the specimen holder to minimize noise, electrical crosstalk, and signal shunting.</li>
	<li>Assemble both halves of the specimen holder with four screws and connect the perfusion line. Dry the electrode in the lower half of the specimen holder and connect the anode cable to the inner retinal side and the cathode cable to the photoreceptor side.</li>
	<li>Perfuse the specimen holder with at least 1 mL/min of oxygenated Ames&#39; medium for 10-20 min to allow time for light responses to stabilize.</li>
</ol>
6. Recording retinal neuronal cell function
<ol>
	<li>Record response families by exposing the retina to light flashes of increasing intensity. For example, record mouse rod photoreceptor response families with light intensities ranging from approximately 10 to 1,000 photons/&#181;m2 and those from mouse ON-bipolar cells with light intensities ranging from approximately 0.6 to 20 photons/&#181;m2.</li>
	<li>Measure photoreceptor light responses (a-wave) in the presence of 100 &#181;M barium chloride, which blocks potassium channels in M&#252;ller glial cells, and 40 &#181;M DL-AP4, which blocks glutamatergic signal transmission to ON-bipolar cells (Figure 2B).</li>
	<li>To isolate the b-wave, which originates from the function of the ON-bipolar cells, first record combined light responses from both photoreceptor and ON-bipolar cells in the presence of barium chloride alone (Figure 2A). Then, perfuse for 5-10 min with Ames&#39; medium containing barium chloride, as well as DL-AP4, and record photoreceptor responses to the same light stimuli as before (Figure 2B). Subtract photoreceptor responses from combined photoreceptor and ON-bipolar cell responses, thus calculating ON-bipolar cell light responses alone (Figure 2C).</li>
</ol>
7. Optimizing ON-bipolar cell function
NOTE: The b-wave, which originates from the ON-bipolar cells, is highly sensitive to the temperature in the specimen holder and the perfusion rate.
<ol>
	<li>Maintain a perfusion rate of at least 0.5 mL/min to obtain the b-wave. 
	NOTE: A higher perfusion rate of 1-2 mL/min is preferable to maintain a large and stable response from ON-bipolar cells.</li>
	<li>Ensure that for a given perfusion rate, the temperature in the specimen holder near the retina is close to body temperature (i.e., approximately 35-38 &#176;C). 
	NOTE: It is important adjust the temperature of the perfusate, which is heated before it reaches the ex vivo ERG specimen holder, so that it is within the optimal temperature range at the retina.</li>
	<li>Store eye cups for later experimentation protected from light and in oxygenated Ames&#39; medium at room temperature to maintain normal a- and b-waves for several hours.</li>
</ol>

<ol>
	<li>Perlman, I., Kolb, H., Fernandez, E., Nelson, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Webvision: The Organization of the Retina and Visual System. , (1995).</li><li>Bonezzi, P. J., Tarchick, M. J., Renna, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ex+vivo+electroretinograms+made+easy:+performing+ERGs+using+3D+printed+components.">Ex vivo electroretinograms made easy: performing ERGs using 3D printed components.</a> Journal of Physiology. 598 (21), 4821-4842 (2020).</li><li>Vinberg, F., Kefalov, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simultaneous+ex+vivo+functional+testing+of+two+retinas+by+in+vivo+electroretinogram+system.">Simultaneous ex vivo functional testing of two retinas by in vivo electroretinogram system.</a> Journal of Visualized Experiments. (99), e52855 (2015).</li><li>Heckenlively, J. R., Arden, G. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Principles and Practice of Clinical Electrophysiology of Vision. , (2006).</li><li>Vinberg, F., Kolesnikov, A. V., Kefalov, V. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ex+vivo+ERG+analysis+of+photoreceptors+using+an+in+vivo+ERG+system.">Ex vivo ERG analysis of photoreceptors using an in vivo ERG system.</a> Vision Research. 101, 108-117 (2014).</li><li>Winkler, B. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium+and+the+fast+and+slow+components+of+P3+of+the+electroretinogram+of+the+isolated+rat+retina.">Calcium and the fast and slow components of P3 of the electroretinogram of the isolated rat retina.</a> Vision Research. 14 (1), 9-15 (1974).</li><li>Green, D. G., Kapousta-Bruneau, N. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+dissection+of+the+electroretinogram+from+the+isolated+rat+retina+with+microelectrodes+and+drugs.">A dissection of the electroretinogram from the isolated rat retina with microelectrodes and drugs.</a> Visual Neuroscience. 16 (4), 727-741 (1999).</li><li>Abbas, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Revival+of+light+signalling+in+the+postmortem+mouse+and+human+retina.">Revival of light signalling in the postmortem mouse and human retina.</a> Nature. , (2022).</li><li>Becker, S., Carroll, L. S., Vinberg, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diabetic+photoreceptors:+Mechanisms+underlying+changes+in+structure+and+function.">Diabetic photoreceptors: Mechanisms underlying changes in structure and function.</a> Visual Neuroscience. 37, (2020).</li><li>Kantola, L., Piccolino, M., Wade, N. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+action+of+light+on+the+retina:+Translation+and+commentary+of+Holmgren+(1866).">The action of light on the retina: Translation and commentary of Holmgren (1866).</a> Journal of the History of the Neurosciences. 28 (4), 399-415 (2019).</li><li>Frank, R. N., Dowling, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rhodopsin+photoproducts:+effects+on+electroretinogram+sensitivity+in+isolated+perfused+rat+retina.">Rhodopsin photoproducts: effects on electroretinogram sensitivity in isolated perfused rat retina.</a> Science. 161 (3840), 487-489 (1968).</li><li>Hamasaki, D. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effect+of+sodium+ion+concentration+on+the+electroretinogram+of+the+isolated+retina+of+the+frog.">The effect of sodium ion concentration on the electroretinogram of the isolated retina of the frog.</a> Journal of Physiology. 167 (1), 156-168 (1963).</li><li>Granit, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+components+of+the+retinal+action+potential+in+mammals+and+their+relation+to+the+discharge+in+the+optic+nerve.">The components of the retinal action potential in mammals and their relation to the discharge in the optic nerve.</a> Journal of Physiology. 77 (3), 207-239 (1933).</li><li>Donner, K., Hemila, S., Koskelainen, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Temperature-dependence+of+rod+photoresponses+from+the+aspartate-treated+retina+of+the+frog+(Rana+temporaria).">Temperature-dependence of rod photoresponses from the aspartate-treated retina of the frog (Rana temporaria).</a> Acta Physiologica Scandinavica. 134 (4), 535-541 (1988).</li><li>Green, D. G., Kapousta-Bruneau, N. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrophysiological+properties+of+a+new+isolated+rat+retina+preparation.">Electrophysiological properties of a new isolated rat retina preparation.</a> Vision Research. 39 (13), 2165-2177 (1999).</li><li>Luke, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+isolated+perfused+bovine+retina--a+sensitive+tool+for+pharmacological+research+on+retinal+function.">The isolated perfused bovine retina--a sensitive tool for pharmacological research on retinal function.</a> Brain Research Protocols. 16 (1-3), 27-36 (2005).</li><li>Becker, S., Carroll, L. S., Vinberg, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rod+phototransduction+and+light+signal+transmission+during+type+2+diabetes.">Rod phototransduction and light signal transmission during type 2 diabetes.</a> BMJ Open Diabetes Research and Care. 8 (1), 001571 (2020).</li><li>Nymark, S., Haldin, C., Tenhu, H., Koskelainen, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+method+for+measuring+free+drug+concentration:+retinal+tissue+as+a+biosensor.">A new method for measuring free drug concentration: retinal tissue as a biosensor.</a> Investigative Ophthalmology &amp; Visual Science. 47 (6), 2583-2588 (2006).</li><li>Winkler, B. S., Kapousta-Bruneau, N., Arnold, M. J., Green, D. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+inhibiting+glutamine+synthetase+and+blocking+glutamate+uptake+on+b-wave+generation+in+the+isolated+rat+retina.">Effects of inhibiting glutamine synthetase and blocking glutamate uptake on b-wave generation in the isolated rat retina.</a> Visual Neuroscience. 16 (2), 345-353 (1999).</li></ol>

None of the authors has any conflicts of interest to disclose.

Originally developed in 1865 by Holmgren to measure retinal light responses from the amphibian retina10, technical constraints initially prevented the ERG from being widely used. Nevertheless, seminal studies by Ragnar Granit and others identified the cellular origins of the ERG and measured photoreceptor and ON-bipolar cell responses ex vivo11,12,13. Since then, refined methods have allowed more...

Electroretinography measures retinal function in response to light1. It is integral to studying retinal physiology and pathophysiology, and measuring the success of therapies for retinal diseases. The in vivo ERG is widely used to assess retinal function in intact organisms, but it has significant limitations2,3. Amongst these, the quantitative analysis of individual retinal cell types in the in vivo ERG is hampered, since it records the sum of potential changes, and therefore overlaying responses, from all retinal cells to light stimuli4. Furthermore, it does not readily allow addition of drugs to the retina, is vulnerable to systemic influences, and has a relatively low signal-to-noise ratio. These disadvantages are eliminated in the ex vivo ERG that investigates the function of the isolated retina2,3,5,6. The ex vivo ERG allows the recording of large and stable responses from specific retinal cell types by addition of pharmacological inhibitors and easy evaluation of therapeutic agents, which can be added to the superfusate. At the same time, it removes influences of systemic effects and eliminates physiological noise (e.g., heartbeat or breathing).
In the ex vivo ERG, retinas or retinal samples are isolated and mounted photoreceptor-side up on the dome of the specimen holder3,5. The specimen holder is assembled, connected to a perfusion system that supplies the retina with heated, oxygenated media, and placed onto the stage of a microscope, which has been modified to deliver computer-controlled light stimuli. To record the responses elicited by light, the specimen holder is connected to an amplifier, digitizer, and recording system (Figure 1). This technique allows isolation of responses from rod and cone photoreceptors, ON-bipolar cells, and M&#252;ller glia by changing the parameters of the light stimuli and adding pharmacological agents.
An existing patch clamp or multi-electrode array (MEA) setup can be converted to record ex vivo ERG, either in conjunction with a commercially available ex vivo ERG adapter or a custom polycarbonate computer numerical control (CNC)-machined specimen holder, to measure light responses in retinas from small animal models, such as mice. This modification increases the accessibility of ex vivo ERG while minimizing the need for specialized equipment. The design of the specimen holder simplifies the mounting technique and integrates electrodes, eliminating the need for manipulation of microelectrodes compared to previously reported transretinal ex vivo ERG methods7. The perfusion rate and temperature inside the specimen holder are important factors that affect the response properties from photoreceptors and ON-bipolar cells. By adjusting these conditions, the ex vivo ERG can be reliably recorded from the isolated mouse retina over prolonged periods of time. Optimized experimental conditions allow ex vivo ERG recordings in retinal punches from larger retinas, including large animal eyes and human donor eyes8.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>2 mm socket</td>
			<td>WPI</td>
			<td>2026-10</td>
			<td>materials to prepare electrode</td>
		</tr>
		<tr>
			<td>Ag/AgCl Electrode</td>
			<td>World Precision Instruments</td>
			<td>EP1</td>
			<td>materials to prepare electrode</td>
		</tr>
		<tr>
			<td>Ames&#39; medium</td>
			<td>Sigma Aldrich</td>
			<td>A1420</td>
			<td>perfusion media</td>
		</tr>
		<tr>
			<td>barium chloride</td>
			<td>Sigma Aldrich</td>
			<td>B0750</td>
			<td>potassium channel blocker</td>
		</tr>
		<tr>
			<td>DL-AP4</td>
			<td>Tocris</td>
			<td>0101</td>
			<td>broad spectrum glutamatergic antagonist</td>
		</tr>
		<tr>
			<td>OcuScience Ex Vivo ERG Adapter</td>
			<td>OcuScience</td>
			<td>n/a</td>
			<td>ex vivo ERG specimen holder</td>
		</tr>
		<tr>
			<td>Threaded luer connector</td>
			<td>McMaster-Carr</td>
			<td>51525K222 or 51525K223</td>
			<td>materials to prepare electrode</td>
		</tr>
	</tbody>
</table>

optimizing the setup and conditions for ex vivo electroretinogram to study retina function in small and large eyes

Measurements of retinal neuronal light responses are critical to investigating the physiology of the healthy retina, determining pathological changes in retinal diseases, and testing therapeutic interventions. The ex vivo electroretinogram (ERG) allows the quantification of contributions from individual cell types in the isolated retina by addition of specific pharmacological agents and evaluation of tissue-intrinsic changes independently of systemic influences. Retinal light responses can be measured using a specialized ex vivo ERG specimen holder and recording setup, modified from existing patch clamp or microelectrode array equipment. Particularly, the study of ON-bipolar cells, but also of photoreceptors, has been hampered by the slow but progressive decline of light responses in the ex vivo ERG over time. Increased perfusion speed and adjustment of the perfusate temperature improve ex vivo retinal function and maximize response amplitude and stability. The ex vivo ERG uniquely allows the study of individual retinal neuronal cell types. In addition, improvements to maximize response amplitudes and stability allow the investigation of light responses in retina samples from large animals, as well as human donor eyes, making the ex vivo ERG a valuable addition to the repertoire of techniques used to investigate retinal function.

Ex vivo ERG enables recording of reproducible and stable photoreceptor and ON-bipolar cell light responses, for example, from the mouse retina (Figure 2A-C). Recording of photoreceptor responses from human donor retinas is possible with up to 5 h postmortem delay of enucleation (Figure 2D) and of ON-bipolar cell responses with a &#60;20 min enucleation delay (Figure 2E). Important parameter...

Watch this Scientific Journal Video about Optimizing the Setup and Conditions for Ex Vivo Electroretinogram to Study Retina Function in Small and Large Eyes at JoVE.com

Optimizing the Setup and Conditions for Ex Vivo Electroretinogram to Study Retina Function in Small and Large Eyes

Modification of existing multielectrode array or patch clamp equipment makes the ex vivo electroretinogram more widely accessible. Improved methods to record and maintain ex vivo light responses facilitate the study of photoreceptor and ON-bipolar cell function in the healthy retina, animal models of eye diseases, and human donor retinas.

Optimizing the Setup and Conditions for Ex Vivo Electroretinogram to Study Retina Function in Small and Large Eyes

Measurements of retinal neuronal light responses are critical to investigating the physiology of the healthy retina, determining pathological changes in ...

optimizing-setup-conditions-for-ex-vivo-electroretinogram-to-study

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

2.2K Views.  University of Utah. Modification of existing multielectrode array or patch clamp equipment makes the ex vivo electroretinogram more widely accessible. Improved methods to record and maintain ex vivo light responses facilitate the study of photoreceptor and ON-bipolar cell function in the healthy retina, animal models of eye diseases, and human donor retinas.

Video: Optimizing the Setup and Conditions for Ex Vivo Electroretinogram to Study Retina Function in Small and Large Eyes

In vivo Electroporation of Developing Mouse Retina

The functional characterization of genes expressed during mammalian retinal development remains a significant challenge. Gene targeting to generate constitutive or conditional loss of function knockouts remains cost and labor intensive, as well as time consuming. Adding to these challenges, retina expressed genes may have essential roles outside the retina leading to unintended confounds when using a knockout approach. Furthermore, the ability to ectopically express a gene in a gain of function experiment can be extremely valuable when attempting to identify a role in cell fate specification and/or terminal differentiation.
We present a method for the rapid and efficient incorporation of DNA plasmids into the neonatal mouse retina by electroporation. The application of short electrical impulses above a certain field strength results in a transient increase in plasma membrane permeability, facilitating the transfer of material across the membrane 1,2,3,4. Groundbreaking work demonstrated that electroporation could be utilized as a method of gene transfer into mammalian cells by inducing the formation of hydrophilic plasma membrane pores allowing the passage of highly charged DNA through the lipid bilayer 5. Continuous technical development has resulted in the viability of electroporation as a method for in vivo gene transfer in multiple mouse tissues including the retina, the method for which is described herein 6, 7, 8, 9, 10. 
DNA solution is injected into the subretinal space so that DNA is placed between the retinal pigmented epithelium and retina of the neonatal (P0) mouse and electrical pulses are applied using a tweezer electrode. The lateral placement of the eyes in the mouse allows for the easy orientation of the tweezer electrode to the necessary negative pole-DNA-retina-positive pole alignment. Extensive incorporation and expression of transferred genes can be identified by postnatal day 2 (P2). Due to the lack of significant lateral migration of cells in the retina, electroporated and non-electroporated regions are generated. Non-electroporated regions may serve as internal histological controls where appropriate. 
Retinal electroporation can be used to express a gene under a ubiquitous promoter, such as CAG, or to disrupt gene function using shRNA constructs or Cre-recombinase. More targeted expression can be achieved by designing constructs with cell specific gene promoters. Visualization of electroporated cells is achieved using bicistronic constructs expressing GFP or by co-electroporating a GFP expression construct. Furthermore, multiple constructs may be electroporated for the study of combinatorial gene effects or simultaneous gain and loss of function of different genes. Retinal electroporation may also be utilized for the analysis of genomic cis-regulatory elements by generating appropriate expression constructs and deletion mutants. Such experiments can be used to identify cis-regulatory regions sufficient or required for cell specific gene expression 11. Potential experiments are limited only by construct availability.

In vivo Electroporation of Developing Mouse Retina

A method for the incorporation of plasmid DNA into murine retinal cells for the purpose of performing either gain- or loss of function studies in vivo is presented. This method capitalizes on the transient increase in permeability of cell plasma membranes induced by the application of an external electrical field.

The functional characterization of genes expressed during mammalian retinal development remains a significant challenge.  Gene targeting to generate ...

Methods for Study of Neuronal Morphogenesis: Ex vivo RNAi Electroporation in Embryonic Murine Cerebral Cortex

The cerebral cortex directs higher cognitive functions. This six layered structure is generated in an inside-first, outside-last manner, in which the first born neurons remain closer to the ventricle while the last born neurons migrate past the first born neurons towards the surface of the brain1. In addition to neuronal migration2, a key process for normal cortical function is the regulation of neuronal morphogenesis3. While neuronal morphogenesis can be studied in vitro in primary cultures, there is much to be learned from how these processes are regulated in tissue environments.
We describe techniques to analyze neuronal migration and/or morphogenesis in organotypic slices of the cerebral cortex4,6. A pSilencer modified vector is used which contains both a U6 promoter that drives the double stranded hairpin RNA and a separate expression cassette that encodes GFP protein driven by a CMV promoter7-9. Our approach allows for the rapid assessment of defects in neurite outgrowth upon specific knockdown of candidate genes and has been successfully used in a screen for regulators of neurite outgrowth8. Because only a subset of cells will express the RNAi constructs, the organotypic slices allow for a mosaic analysis of the potential phenotypes. Moreover, because this analysis is done in a near approximation of the in vivo environment, it provides a low cost and rapid alternative to the generation of transgenic or knockout animals for genes of unknown cortical function. Finally, in comparison with in vivo electroporation technology, the success of ex vivo electroporation experiments is not dependant upon proficient surgery skill development and can be performed with a shorter training time and skill.

Methods for Study of Neuronal Morphogenesis: Ex vivo RNAi Electroporation in Embryonic Murine Cerebral Cortex

To conduct a rapid assessment of the function of genes in the development of cerebral cortex, we describe methods involving the ex vivo electroporation of plasmids co-expressing inhibitory RNA (RNAi) and GFP in murine embryonic cortex. This protocol is amenable to the study of various aspects of neurodevelopment such as neurogenesis, neuronal migration and neuronal morphogenesis including dendrite and axon outgrowth.

The cerebral cortex directs higher cognitive functions. This six layered structure is generated in an inside-first, outside-last manner, in which the ...

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow for an in-depth characterization to identify pathways involved in these events. Cerebellar granule neurons (CGNs) that are derived from the developing cerebellum are an ideal model system that allows for morphological analyses. Here, we describe a method of how to genetically manipulate CGNs and how to study axono- and dendritogenesis of individual neurons. With this method the effects of RNA interference, overexpression or small molecules can be compared to control neurons. In addition, the rodent cerebellar cortex is an easily accessible in vivo system owing to its predominant postnatal development. We also present an in vivo electroporation technique to genetically manipulate the developing cerebella and describe subsequent cerebellar analyses to assess neuronal morphology and migration.

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Neuronal morphogenesis and migration are crucial events underlying proper brain development. Here, we describe methods to genetically manipulate cultured cerebellar granule neurons and the developing cerebellum for the assessment of morphology and migratory characteristics of neurons.

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow ...

Mouse Hindbrain Ex Vivo Culture to Study Facial Branchiomotor Neuron Migration

Embryonic neurons are born in the ventricular zone of the brain, but subsequently migrate to new destinations to reach appropriate targets. Deciphering the molecular signals that cooperatively guide neuronal migration in the embryonic brain is therefore important to understand how the complex neural networks form which later support postnatal life. Facial branchiomotor (FBM) neurons in the mouse embryo hindbrain migrate from rhombomere (r) 4 caudally to form the paired facial nuclei in the r6-derived region of the hindbrain. Here we provide a detailed protocol for wholemount ex vivo culture of mouse embryo hindbrains suitable to investigate the signaling pathways that regulate FBM migration. In this method, hindbrains of E11.5 mouse embryos are dissected and cultured in an open book preparation on cell culture inserts for 24 hr. During this time, FBM neurons migrate caudally towards r6 and can be exposed to function-blocking antibodies and small molecules in the culture media or heparin beads loaded with recombinant proteins to examine roles for signaling pathways implicated in guiding neuronal migration.

Mouse Hindbrain Ex Vivo Culture to Study Facial Branchiomotor Neuron Migration

Embryonic neurons are born in the ventricular zone of the neural tube, but migrate to reach appropriate targets. Facial branchiomotor (FBM) neurons are a useful model to study neuronal migration. This protocol describes the wholemount ex vivo culture of mouse embryo hindbrains to investigate mechanisms that regulate FBM migration.

Embryonic neurons are born in the ventricular zone of the brain, but subsequently migrate to new destinations to reach appropriate targets. Deciphering ...

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first neural circuit in the AOS pathway, called the accessory olfactory bulb (AOB), plays an important role in establishing sex-typical behaviors such as territorial aggression and mating. This small (&#60;1 mm3) circuit possesses the capacity to distinguish unique behavioral states, such as sex, strain, and stress from chemosensory cues in the secretions and excretions of conspecifics. While the compact organization of this system presents unique opportunities for recording from large portions of the circuit simultaneously, investigation of sensory processing in the AOB remains challenging, largely due to its experimentally disadvantageous location in the brain. Here, we demonstrate a multi-stage dissection that removes the intact AOB inside a single hemisphere of the anterior mouse skull, leaving connections to both the peripheral vomeronasal sensory neurons (VSNs) and local neuronal circuitry intact. The procedure exposes the AOB surface to direct visual inspection, facilitating electrophysiological and optical recordings from AOB circuit elements in the absence of anesthetics. Upon inserting a thin cannula into the vomeronasal organ (VNO), which houses the VSNs, one can directly expose the periphery to social odors and pheromones while recording downstream activity in the AOB. This procedure enables controlled inquiries into AOS information processing, which can shed light on mechanisms linking pheromone exposure to changes in behavior.

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory bulb (AOB) has been difficult to study in the context of sensory coding. Here, we demonstrate a dissection that produces an ex vivo preparation in which AOB neurons remain functionally connected to their peripheral inputs, facilitating research into information processing of mouse pheromones and kairomones.

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first ...

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of the olfactory epithelium continuously give rise to new sensory neurons that extend their axons into the olfactory bulb, where they face the challenge to integrate into existing circuitry. Because of this particular feature, the olfactory system represents a unique opportunity to monitor axonal wiring and guidance, and to investigate synapse formation. Here we describe a procedure for in vivo labeling of sensory neurons and subsequent visualization of axons in the olfactory system of larvae of the amphibian Xenopus laevis. To stain sensory neurons in the olfactory organ we adopt the electroporation technique. In vivo electroporation is an established technique for delivering fluorophore-coupled dextrans or other macromolecules into living cells. Stained sensory neurons and their axonal processes can then be monitored in the living animal either using confocal laser-scanning or multiphoton microscopy. By reducing the number of labeled cells to few or single cells per animal, single axons can be tracked into the olfactory bulb and their morphological changes can be monitored over weeks by conducting series of in vivo time lapse imaging experiments. While the described protocol exemplifies the labeling and monitoring of olfactory sensory neurons, it can also be adopted to other cell types within the olfactory and other systems.

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

We describe a protocol for in vivo labeling of olfactory sensory neurons by electroporation and subsequent confocal laser-scanning or multiphoton microscopy to visualize neuronal morphology and its development over time.

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of ...

Using the Electroretinogram to Assess Function in the Rodent Retina and the Protective Effects of Remote Limb Ischemic Preconditioning

The ERG is the sum of all retinal activity. The ERG is usually recorded from the cornea, which acts as an antenna that collects and sums signals from the retina. The ERG is a sensitive measure of changes in retinal function that are pan-retinal, but is less effective for detecting damage confined to a small area of retina. In the present work we describe how to record the &#8216;flash&#8217; ERG, which is the potential generated when the retina is exposed to a brief light flash. We describe methods of anaesthesia, mydriasis and corneal management during recording; how to keep the retina dark adapted; electrode materials and placement; the range and calibration of stimulus energy; recording parameters and the extraction of data. We also describe a method of inducing ischemia in one limb, and how to use the ERG to assess the effects of this remote-from-the-retina ischemia on retinal function after light damage. A two-flash protocol is described which allows isolation of the cone-driven component of the dark-adapted ERG, and thereby the separation of the rod and cone components. Because it can be recorded with techniques that are minimally invasive, the ERG has been widely used in studies of the physiology, pharmacology and toxicology of the retina. We describe one example of this usefulness, in which the ERG is used to assess the function of the light-damaged retina, with and without a neuroprotective intervention; preconditioning by remote ischemia.

The electroretinogram (ERG) is an electrical potential generated by the retina in response to light. This paper describes how to use the ERG to assess retinal function, in dark-adapted rats, and how it can be can be used to assess a neuroprotective intervention, in the present case remote ischemic preconditioning.

The ERG is the sum of all retinal activity. The ERG is usually recorded from the cornea, which acts as an antenna that collects and sums signals from the ...

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise from extra-retinal sources. Ex vivo ERG provides an efficient method to dissect the function of retinal cells directly from an intact isolated retina of animals or donor eyes. In addition, ex vivo ERG can be used to test the efficacy and safety of potential therapeutic agents on retina tissue from animals or humans. We show here how commercially available in vivo ERG systems can be used to conduct ex vivo ERG recordings from isolated mouse retinas. We combine the light stimulation, electronic and heating units of a standard in vivo system with custom-designed specimen holder, gravity-controlled perfusion system and electromagnetic noise shielding to record low-noise ex vivo ERG signals simultaneously from two retinas with the acquisition software included in commercial in vivo systems. Further, we demonstrate how to use this method in combination with pharmacological treatments that remove specific ERG components in order to dissect the function of certain retinal cell types.

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Ex vivo ERG can be used to record electrical activity of retinal cells directly from isolated intact retinas of animals or humans. We demonstrate here how common in vivo ERG systems can be adapted for ex vivo ERG recordings in order to dissect the electrical activity of retinal cells.

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise ...

Rabies Necropsy Techniques in Large and Small Animals

The New York State Department of Health (NYSDOH) Rabies Laboratory receives between 6,000 to 9,000 specimens annually and performs rabies testing for the entire state, with the exception of New York City. The Rabies laboratory necropsies a variety of animals ranging in size from bats to bovids. Most of these specimens are animals exhibiting neurological signs, however, less than 10% actually test positive for rabies; implying trauma, lesions or other infectious agents as the cause of these symptoms. Due to the risk of aerosolizing undiagnosed infectious agents, the Rabies Laboratory does not use power tools or saws. Three necropsy techniques will be presented for animals whose skulls are impenetrable with scissors. The laboratory has implemented these techniques to decrease potential exposure to infectious agents, eliminate unnecessary manipulation of the specimen and reduce processing time. The advantages of a preferred technique opposed to another is subject to the trained individual processing the specimen.

The goal of this protocol is to demonstrate safe necropsy techniques in small and large animals to obtain satisfactory tissue samples for rabies testing.

The New York State Department of Health (NYSDOH) Rabies Laboratory receives between 6,000 to 9,000 specimens annually and performs rabies testing for the ...

Transpupillary Two-Photon In Vivo Imaging of the Mouse Retina

The retina transforms light signals from the environment into electrical signals that are propagated to the brain. Diseases of the retina are prevalent and cause visual impairment and blindness. Understanding how such diseases progress is critical to formulating new treatments. In vivo&#160;microscopy in animal models of disease is a powerful tool for understanding neurodegeneration and has led to important progress towards treatments of conditions ranging from Alzheimer&#39;s disease to stroke. Given that the retina is the only central nervous system structure inherently accessible by optical approaches, it naturally lends itself towards in vivo imaging. However, the native optics of the lens and cornea present some challenges for effective imaging access.
This protocol outlines methods for in vivo two-photon imaging of cellular cohorts and structures in the mouse retina at cellular resolution, applicable for both acute- and chronic-duration imaging experiments. It presents examples of retinal ganglion cell (RGC), amacrine cell, microglial, and vascular imaging using a suite of labeling techniques including adeno-associated virus (AAV) vectors, transgenic mice, and inorganic dyes. Importantly, these techniques extend to all cell types of the retina, and suggested methods for accessing other cellular populations of interest are described. Also detailed are example strategies for manual image postprocessing for display and quantification. These techniques are directly applicable to studies of retinal function in health and disease.

In vivo imaging is a powerful tool for the study of biology in health and disease. This protocol describes transpupillary imaging of the mouse retina with a standard two-photon microscope. It also demonstrates different in vivoimaging methods to fluorescently label multiple cellular cohorts of the retina.

The retina transforms light signals from the environment into electrical signals that are propagated to the brain. Diseases of the retina are prevalent ...

Optimizing the Setup and Conditions for Ex Vivo Electroretinogram to Study Retina Function in Small and Large Eyes

Podsumowanie

Przeglądaj więcej filmów

Rozdziały w tym wideo

In vivo Electroporation of Developing Mouse Retina

Methods for Study of Neuronal Morphogenesis: Ex vivo RNAi Electroporation in Embryonic Murine Cerebral Cortex

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Mouse Hindbrain Ex Vivo Culture to Study Facial Branchiomotor Neuron Migration

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

Using the Electroretinogram to Assess Function in the Rodent Retina and the Protective Effects of Remote Limb Ischemic Preconditioning

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Rabies Necropsy Techniques in Large and Small Animals

Transpupillary Two-Photon In Vivo Imaging of the Mouse Retina